Download Ecto-enzymes ofmammary gland and its tumours

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Biochem. J. (1980) 191,45-51
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Ecto-enzymes of mammary gland and its tumours
Ca2+- OR Mg2+-STIMULATED ADENOSINE TRIPHOSPHATASE AND ITS PERTURBATION BY
CONCANAVALIN A
Coralie A. Carothers CARRAWAY, Frank J. CORRADO IV, Douglas D. FOGLE
and Kermit L. CARRAWAY
Department ofBiochemistry, Oklahoma State University, Stillwater, OK 74078, U.S.A.
(Received 18 December 1979/Accepted 13 May 1980)
Intact viable 13762 mammary-adenocarcinoma ascites cells hydrolyse added ATP. The
localization of hydrolysis product and inactivation by the slowly penetrating chemical
reagent diazotized sulphanilic acid indicate that this ATPase is at the external surface of
the cell. A number of features differentiate this enzyme from mitochondrial, myosin and
cation-transport ATPases. It is stimulated by either Ca2+ or Mg2+ and has little or no
activity in their absence. It is insensitive to ouabain, oligomycin and azide. It is the
major ATPase activity found in homogenates of gently disrupted 13762 cells. The
ATPase activity is inhibited at high substrate concentrations and shows an apparent
stimulation by concanavalin A in isolated membranes, but not in intact cells. The
stimulation by concanavalin A results predominantly from a release from substrate
inhibition.
Cell-surface enzymes are potentially useful
markers of membrane changes which occur as a
result of cell differentiation, various disease states or
artificial perturbations (Stefanovic et al., 1976;
Carraway et al., 1976, 1979). A problem with the
use of marker enzymes is the lack of information on
the properties of these enzymes and their
physiological functions. For example, an ectoATPase activity (Trams & Lauter, 1974; DePierre
& Karnovsky, 1974a,b; Ronquist & Agren, 1975;
Stefanovic et al., 1976) has been described in a
number of cell types. Its external location suggests
that it is different from previously described
ATPases, which are found at the interior surface of
plasma membranes and often supply the energy
requirement for transport functions. In the human
erythrocyte membrane, the best-studied membrane
model, there are at least three ATPases (Drickamer,
1975). The ouabain-sensitive enzyme is involved in
Na+ and K+ transport and requires Mg2+. A second
enzyme is involved in Ca2+ transport and requires
Mg2+ for activity. The third enzyme is designated
Mg2+-ATPase, and no function has been ascribed to
it. In some more complex cells an ATPase activity
has been described which is stimulated by either
Mg2+ or Ca2+ (Parkinson & Radde, 1971). Ehrlich
ascites-tumour cells (Ronquist & Agren, 1975) and
some neural cells (Stefanovic et al., 1976) hydrolyse
exogenously added ATP by such an enzyme. Several
Vol. 191
previous studies have established the existence of
ecto-ATPases which are stimulated by Mg2+
(DePierre & Karnovsky, 1974a,b; Trams & Lauter,
1974). Whether these activities are also stimulated
by Ca2+ has not been reported.
In the present study we have used established
criteria (DePierre & Karnovsky, 1974a) to
demonstrate the presence at the cell surface of 13762
mammary-adenocarcinoma cells of an ATPase
activity which is stimulated by either Mg2+ or Ca2+.
The enzyme in isolated membranes shows an
apparent activation on treatment with concanavalin
A. However, this activation is due largely to release
from substrate inhibition at high ATP
concentrations. Intact cells show neither
concanavalin A activation nor substrate inhibition.
Experimental
Materials
ATP,
GTP,
ADP,
glucose
6-phosphate,
oligomycin, iodoacetic acid and ouabain were
obtained from Sigma, as were buffers and salts, all
reagent grade. p-Nitrophenyl phosphate was
supplied by Nutritional Biochemicals Corp.,
sulphanilic acid by Fisher Scientific Co. and
concanavalin A by Miles Laboratories [y-32PIATP
was from Amersham.
0306-3275/80/100045-07$01.50/1 (© 1980 The Biochemical Society
46
C. A. C. Carraway, F. J. Corrado IV, D. D. Fogle and K. L. Carraway
Isolation of cells and membranes
The 13762 MAT-A and MAT-C l mammary
ascites adenocarcinomas were maintained in Fischer
344-strain female rats. Cells and Zn2+-stabilized
membrane envelopes of the cells were obtained as
previously described (Carraway et al., 1976, 1979).
Membranes were stored frozen for no longer than 3
days before enzyme studies and were frozen and
thawed no more than once, since the ATPase
appears somewhat labile.
Enzyme assays
Ca2+- or Mg2+-activated ATPase was assayed by
incubating cells (approx. 106 per assay) or
membranes (approx. 50,g of protein) at 37°C for
2-10min in 25 mM-histidine/25 mM-imidazole/
120 mM-KCI/1 mM-ouabain (pH 8.2) and the appropriate concentration of ATP and Ca2+ or Mg2+
(usually 5mM bivalent cation) in a 0.4 ml assay
volume. Both unlabelled ATP and [y-32P]ATP were
used as substrates. The reaction was terminated by
addition of 0.5 ml of 10% (w/v) trichloroacetic acid.
Samples chilled on ice for 10min were centrifuged
and samples taken for assay of P1 by the method of
Chen et al. (1956) or by scintillation counting after
treatment with charcoal to remove nucleotides (Doss
et al., 1979). Controls containing no enzyme were
used to correct for non-enzymic hydrolysis. The
same procedure was used in assays with GTP, ADP,
glucose 6-phosphate and p-nitrophenyl phosphate as
substrate, by using unlabelled compounds and
chemical analyses. Ouabain-sensitive ATPase (Shin
& Carraway, 1973), lactate dehydrogenase
(Neilands, 1955) and 5'-nucleotidase (Carraway et
al., 1976) were assayed as previously described.
Other assays
Protein was measured by the procedure of Lowry
et al. (1951). Cells were quantified by counting in a
haemocytometer.
Treatments with diazotized sulphanilic acid
Diazotized sulphanilic acid was prepared by the
procedure of DePierre & Karnovsky (1974a).
Treatments with it before and after cell
homogenization were performed as follows.
(1) Treatment of intact cells. Washed intact cells
(5.3 x 107) were treated for 15 min at 370C with
1.0mM- or 5.0 mM-diazotized sulphanilic acid in
2.0 ml of iso-osmotic (10mM) Hepes [4-(2)-hydroxy-
ethyl)- 1-piperazine-ethanesulphonic acid] /saline
buffer, pH 7.4. The treated cells were washed twice
in Hepes; samples were taken for assays of Ca2+- or
Mg2+-ATPase and lactate dehydrogenase. Another
sample of treated cells was homogenized at 0°C in
40mM-Tris/HCI, pH 7.4, by five strokes of a Dounce
B homogenizer. Over 99% of the cells were broken
by this procedure. The homogenate was assayed for
Ca2+_ or Mg2+-ATPase and lactate dehydrogenase
(Neilands, 1955).
(2) Treatment of homogenate. Washed intact cells
were homogenized in the same manner and samples
of the homogenate subsequently treated with
1.OmM- and 5.0mM-diazotized sulphanilic acid. The
treated homogenates were assayed for Ca2+_ or
Mg2+-ATPase and lactate dehydrogenase.
Results
Cell-surface localization ofA TPase
To evaluate the relationship between the ATPase
and other cell-surface properties, two different
ascites sublines of the 13762 rat mammary
adenocarcinoma, MAT-A and MAT-C 1, were used.
As noted previously (Carraway et al., 1979), these
sublines differ markedly in morphology and
concanavalin A-receptor mobility. However, no
significant differences between the sublines were
found in the properties of the ATPase. Similar
results were obtained previously for 5'-nucleotidase
(Carraway et al., 1979). The experimental values
reported here are averages of duplicate determinations, except as noted, varying less than 15%
from the reported value. The major source of
variation was in the cell and membrane preparations,
resulting in different specific activities. Moreover,
each type of experiment has been repeated on the
other subline for verification of the result. Therefore
the presentation below will simply refer to the cells
as 13762 MAT cells without regard to the subline
used.
To establish that the ATPase of the 13762 cells is
an ecto-enzyme, we have used criteria developed by
DePierre & Karnovsky (1974a). In addition, all
ATPase-perturbation studies were performed in the
presence of Mg2+ and in the presence of Ca2+ in
parallel experiments to show that this enzyme can be
activated by either cation. A number of experimental
results indicate the external membrane location of
the enzyme. (1) Externally added ATP is hydrolysed
by cell populations containing no more than 5%
non-viable cells, as measured by Trypan Blue
exclusion. This evidence does not rule out the
possibility of transient or localized ATP uptake by
the cells. However, such an uptake mechanism, if
present, does not appear to be energy-dependent,
since treatment with azide or iodoacetate as energy
poisons does not significantly decrease ATP hydrolysis by the intact cells. (2) Assays of medium in
which cells have been incubated show less than 3%
of the activity in cells, indicating that ATP
hydrolysis is not due to a released enzyme. (3)
[32P]ATP was hydrolysed by cells that had been
previously loaded with [33P]phosphate (DePierre &
Kamovsky, 1974a). The distribution of 32P and 33P
1980
Mammary-tumour ecto-ATPase
in the cells and supernatant indicated that over 90%
of the enzyme activity must be extracellular. (4) The
ATPase determined in the intact cell represents more
than 90% of the ouabain-insensitive, Ca2+-stimulated, and more than 80% of the ouabain-insensitive, Mg2+-stimulated activity of homogenates
obtained by gentle homogenization of the cells.
Membrane preparations were assayed for Na+- and
K+-dependent ATPase (Shin & Carraway, 1973) in
the presence and absence of ouabain. The ouabaininsensitive ATPase activity represented 60-70% of
the total ATPase activity. These results suggest that
the enzyme studied in the intact cell is the same as
that of the isolated cell-surface membrane and a
major contributor to the total ATPase activity of the
cells. (5) Cells and homogenates were treated with
diazotized sulphanilic acid (DePierre & Karnovsky,
1974a), a slowly penetrating chemical reagent which
reacts covalently with a number of protein functional
groups. Brief treatments of intact 13762 cells with
diazotized sulphanilic acid caused marked decreases
in the ATPase activity, assayed in either intact cells
or homogenates (Table 1). Inactivation of Ca2+- and
Mg2+-stimulated activities was essentially the same.
The diazotized sulphanilic acid treatment of intact
cells had much smaller effects on lactate
dehydrogenase, assayed in homogenates. When
homogenates were treated with diazotized
sulphanilic acid, lactate dehydrogenase was inhibited
by over 95%. ATPase activities in homogenates
and in intact cells treated with diazotized sulphanilic
acid were inhibited almost to the same extent.
The results of the chemical modification experiments closely parallel those of DePierre &
Karnovsky (1974a) on ecto-phosphohydrolases and
strongly support the postulate of an external location
of the ATPase.
47
Kinetic properties and inhibitor effects on A TPase
activities in intact cells and isolated membrane
envelopes
There are several features which distinguish the
activities reported here from the more commonly
studied ATPases, such as those of the erythrocyte
membrane, sarcoplasmic reticulum or mitochondria.
The ecto-ATPase is stimulated independently by
Ca2+ or Mg2+ in the absence of the other cation
either in the intact cell (Fig. 1) or in isolated
membranes (Fig. 2). These membranes are obtained
after Zn2+ 'stabilization' as envelopes ('ghosts')
derived from the cell surface. Phase-contrast
microscopy shows a tear in the envelope through
which the nucleus and cytoplasm were extruded.
Thus both inner and outer surfaces of the membrane
u)
60
O
50
& 40
0
E 30
, 20
C.)
cis
c)
a.
wt
10
un
1.0
2.0
3.0
4.0
[Bivalent cation] (mM)
Fig. 1. Activation of intact 13762-cell A TPase by bivalent
cations
Assays contained 2.5 x 10o cells and 2.5 mM-ATP.
For details see the text. Additions: 0, Ca2+; 0,
Mg2+; U, EDTA, O, no bivalent cations.
Table 1. Effect of diazotized sulphanilic acid (DSA) on A TPase and lactate dehydrogenase activities of whole cells
and homogenates
For details see the text.
ATPase (,umol/h per 101 cells)
A&+ Lactate dehydrogenase
Ca2+
(umol/h per 107 cells)
Mg2+
Treatment
Intact
3.7
55
59
Untreated
4.4
21
23
1 mM-DSA
2
4.8
3
5 mM-DSA
Cells homogenized before DSA treatment
53
690
51
Untreated
410
12
14
1 mM-DSA
27
5mM-DSA
Cells homogenized after DSA treatment
19
610
15
I mM-DSA
7
510
3
5mM-DSA
Vol. 191
C. A. C. Carraway, F. J. Corrado IV, D. D. Fogle and K. L. Carraway
48
Table 2. A TPase and S'-nucleotidase activities in cellfractions during plasma-membrane puriflcation
Values in parentheses represent the degree of purification.
Activity (umol/h per mg of protein)
Mg2+-ATPase
Fraction
Intact cells
Crude membrane pellet
Purified membranes
- - - - -~
- ~~~.
60
._
U
50
.-4v
40
>
L-
._
o
0
4C)
a
9.5
7.8
48 (5.1)
30
20
10
::-
0
1.0
2.0
4.0
3.0
5.0
[Bivalent cation] (mb
Fig. 2. Activation of 13762-cell membrrane A TPase by
bivalent cations
Assays contained 85,ug of membrane Iprotein for the
study with Ca2+ (0) and 34,pg for tihe study with
Mg2+ (0). ATP concentration was 2 mlM.
7
0.250
4
lu
(a)
r-
0.200
c)
0
0.150
U
0.
0
0.100
E
0.050
_.
0
-2
-2 -1
0
2
1/[ATPl (mM
Q0
*
-S
0
2on
C-
;0
E~-.
0
2.5
5.0
6.7
38 (4.8)
5'-Nucleotidase
1.7
1.6
9.9 (5.8)
are accessible. The responses of the activity to Ca2+
and Mg2+ are nearly the same. There is a small
amount of residual activity (approx. 3%) in the
absence of added bivalent cation, which is not
removed by the addition of 1 mM-EDTA. The effects
of the Ca2+ and Mg2+ are additive when they are
added at less than saturating concentrations,
suggesting that they act at the same site.
Oligomycin does not substantially inhibit the
Mg2+- or Ca2+-ATPase activity in intact cells;
homogenate enzyme is inhibited by less than 20%o.
The fact that no substantial increase of ATPase is
observed in breaking the cell permeability barrier in
the preparation of homogenates again suggests that
this ATPase is the major ATPase activity of these
cells. To verify that the membrane and cell-surface
activities are the same enzyme, the purification of
ATPase was compared with 5'-nucleotida$e, ,which
has been established as an ecto-enzyme in these cells
(Carraway et al., 1976). There is a close
correspondence between the purification of the
nucleotidase and Mg2+- or Ca2+-ATPase (Table 2).
The fact that Mg2+- and Ca2+-stimulated activities
show parallel behaviour during membrane isolation
and inhibition by diazotized sulphanilic acid suggests
that they reside on a single enzyme. Kinetiq studies
on the enzyme support this contention. Fig. 3 shows
double-reciprocal plots of enzyme-kinetic data with
ATP as substrate for intact cells and membranes.
Km values are 0.9mm and 0.6mm for the Mg2tactivated enzyme and 1.8mm and 1.1mm for tle
Ca2 -activated enzyme in the intact cells
membranes respectively. In the isolated membranes
substrate inhibition can be seen. This inhibition,
which varies with different membrane preparations,
is greater in the presence of Ca2+ than with Mg2+.
Substrate inhibition has not been observed in intact
cells. No inhibition with increasing ATP concentration is noted at suboptimal bivalent-cation
concentrations (results not shown). Therefore the
substrate inhibition observed is not due to free ATP.
The results suggest that the substrates and inhibitors
for the Ca2+- and Mg2+-activated enzymes are the
respective bivalent-cation-ATP complexes, although
a complete kinetic analysis has not been performed.
Other phosphohydrolase activities
Intact 13762 cells also show substantial
ar*d
6
4
Ca2+-ATPase
8.0
7.5
10.0
1/[ATPI (mM-'
Fig. 3. Lineweaver-Burk plots of inta4 ct 13762-cell (a)
and membrane (b) A TPase
Assays contained 9.2 x 104 cells. For details see the
text. *, Ca2+; 0, Mg2+.
1980
Mammary-tumour ecto-ATPase
49
hydrolytic activity against externally added GTP
under ATPase-assay conditions. Although the Ca2+stimulated activity exhibits simple kinetics (Km for
GTP 3mM), the Mg2+-stimulated GTP hydrolysis
shows a biphasic double-reciprocal plot (Km values
3 and 0.4mM), suggesting two different enzymes.
ADP hydrolysis proceeds at about one-fifth the rate
of ATP hydrolysis and exhibits complex kinetics.
The ATPase activities observed here are not due to
non-specific or other phosphatase activities, since the
isolated membranes prepared by the Zn2+stabilization procedure show essentially no
hydrolytic activities against glucose 6-phosphate and
p-nitrophenyl phosphate. The absence of these
enzyme activities is not due to inactivation of the
enzymes during membrane preparation (Shin &
Caraway, 1973).
Concanavalin A perturbation of the membrane
A TPase
Our previous studies (Carraway et al., 1975)
showed that Mg2+-ATPase of partially purified rat
mammary plasma membranes was activated by the
plant lectin concanavalin A. Similar experiments
with MAT cells and plasma membranes showed
rather unusual behaviour. At low or intermediate
substrate concentrations (up to 2.5 mM-ATP) the
enzyme activity was not substantially changed in
either cells or membranes. Small degrees of either
activation or inhibition (up to about 15%) were
observed in different membrane preparations.
However, at higher ATP concentrations a definite
activation was observed (Table 3). The amount of
activation and the concentration of ATP necessary
to observe activation were variable with different
membrane preparations. Since activation by
concanavalin A and substrate inhibition appeared to
be correlated in different preparations, the kinetics of
the enzyme were examined in the presence and
absence of concanavalin A. Fig. 4 shows that the
increased ATPase activity is not due to a true
activation, but instead results from a relief of
substrate inhibition. As observed previously for the
mammary membranes, the concanavalin A effect
0.20
A
1-1
0.
0
I..
0.16
m
00
E 0.12
0.
.0
o 0.08
E
-
0.04
-2
2
0
4
6
1/[ATP] (mM-')
Fig. 4. Lineweaver-Burk plot of membrane A TPase with
and without concanavalin A
Membranes (43,ug of protein/assay) were incubated
for 30min at 370C in assay medium containing
5 mM-Mg2+ in the absence (A) and presence (M) of
660,ug of concanavalin A/mg of protein (80,ug/ml)
before addition of substrate.
90
,
80
0
-.
E.2
70
0,o
;t 0.
-> .
0-o
60
00
50
0
+50mM-a-MMJ
Table 3. Effect of concanavalin A on membrane
Mg2+-A TPase
Samples were incubated for 30min at 370C with
concanavalin A before enzyme assays. Values are
determinations for a typical membrane preparation.
Conditions
ATP
Concanavalin A
Activity
(mM)
(#g/ml)
(umol/h per mg of protein)
2.5
2.5
2.5
5.0
5.0
5.0
0
200
500
0
200
500
6.9
5.9
6.3
4.2
7.8
10.2
Vol. 191
t
Control
30
0
10
100
1000
[Concanavalin Al (pg/mg of protein)
Fig. 5. Concentration-dependence of concanavalin A
stimulation of membrane A TPase
Membranes (30,ug of protein/assay) were incubated
in assay medium containing 5 mm-Ca2+ and
concanavalin A 15-lOOO,ug/mg of protein (0.489pg/ml)l for 20min at 37°C before assay with
2 mM-ATP. The same experiment performed on
membrane suspensions containing 5mM-Mg2+ gave
a very similar activation plot and identical Hill
coefficient (1.7). Abbreviation: a-MM, a-methyl
mannoside.
50
C. A. C. Carraway, F. J. Corrado IV, D. D. Fogle and K. L. Carraway
(Fig. 5) is concentration-dependent and exhibits
positive co-operativity (Hill coefficient 1.7).
In contrast, no activation of the enzyme was
observed by treatment of intact cells with
concanavalin A at substrate concentrations of
0-5 mM-ATP and concanavalin A concentrations as
high as 500,ug/ml.
Discussion
The ATPase described in this study is unique in a
number of ways. Its exterior location and lack of
response to ouabain, oligomycin and azide
distinguish this enzyme from the Na+,K+- and
Ca2+-ATPases associated with transport of these
ions and from the Mg2+-ATPase present in erythrocytes and other cells. The activities activated by
Ca2+ and by Mg2+ show parallel behaviour under
essentially all conditions tested. The results are best
explained by a single enzyme stimulated by both
cations. The response to cations also distinguishes
this enzyme from other eukaryotic membrane
ATPases and from myosin, which has been
postulated to be present at the cell surface
(Willingham et al., 1974). The behaviour toward
oligomycin, Ca2+ and Mg2+ appears more similar to
some of the bacterial ATPases (Abrams & Smith,
1970) than to the mammalian enzymes. However,
many of the bacterial enzymes are also inhibited by
azide. The ecto-ATPase represents a major ATPhydrolysing activity of the 13762 cells and of their
isolated plasma membranes.
The presence of an ecto-ATPase immediately
raises questions as to its function. An obvious
answer is that it serves to control the extracellular
ATP concentration. It is more difficult to specify
exactly what ATP does at the external surface of
cells and how it gets there. ATP alters cell volume
and ion fluxes (Rorive & Kleinzeller, 1972),
morphology in mast cells (Kruger et al., 1974), cell
adhesion, aggregation and movement in fibroblasts
(Jones, 1966; Knight et al., 1966) and insulin
stimulation of glucose transport in adipocytes
(Chang & Cuatrecasas, 1974). External ATP also
has an effect on p-nitrophenyl phosphate transport
which differs in normal and transformed 3T3 cells
(Rozengurt & Heppel, 1975). The basis or bases for
these ATP effects are unknown, but they may be
related to protein kinase activity at cell surfaces
(Mastro & Rozengurt, 1976). The source of the
extracellular ATP is uncertain, but there is evidence
that ATP may be translocated from the cytosol to
the cell exterior (Trams, 1974).
The observations on substrate inhibition of the
ATPase are intriguing. If substrate inhibition is
explained by a two-site model, the fact that substrate
inhibition is observed in membranes, but not in cells,
may indicate that the second (non-substrate) site is
inside the cell. Such an arrangement could permit
control of the enzyme activity by the cellular ATP
concentration. This control could be over-ridden
from outside the cell by factors such as concanavalin
A, which interact with carbohydrates at the cell
surface. Although the mechanism of action of
concanavalin A is not presently clear, it could cause a
conformational change in the enzyme which blocked
the second ATP site.
Exactly how our results relate to previously
described effects of concanavalin A on membrane
ATPases (Novogrodsky, 1972; Jarett & Smith,
1974; Luly & Emmelot, 1975; Pommier et al., 1975)
is not known. Riordan et al. (1977) have described
an activation of a Mg2+-ATPase of liver membranes
which does not appear to be related to substrate
inhibition and which is sensitive to temperature
effects on the membrane. Whether this is an
ecto-enzyme and is stimulated by Ca2+ as well as by
Mg2+ is unknown. It seems likely that there is more
than one mechanism for ATPase stimulation by
concanavalin A.
Clearly membrane enzymes as sensitive as
ATPases can be altered by a number of membrane
perturbants and should be useful in understanding
membrane structure-function relationships.
The excellent technical assistance of Timothy Chan
and Glendon Jett is gratefully acknowledged. This is
Journal Article J-3310 of the Agricultural Experiment
Station, Oklahoma State University, Stillwater, OK. This
research was conducted in co-operation with the U.S.
Department of Agriculture, Agricultural Research
Service, Southern Region, and supported by the National
Cancer Institute (No-l-CB-33910 and CA 19985) and
the Oklahoma Agricultural Experiment Station.
References
Abrams, A. & Smith, J. B. (1970) Enzymes 3rd Ed. 10,
395-429
Caraway, C. A. C., Jett, G. & Carraway, K. L. (1975)
Biochem. Biophys. Res. Commun. 67, 1301-1306
Carraway, K. L., Fogle, D. D., Chesnut, R. W., Huggins,
J. W. & Carraway, C. A. C. (1976) J. Biol. Chem. 25 1,
6173-6178
Carraway, K. L., Doss, R. C., Huggins, J. W., Chesnut,
R. W. & Carraway, C. A. C. (1979) J. Cell Biol. 83,
529-543
Chang, K.-J. & Cuatrecasas, P. (1974) J. Biol. Chem.
249, 3170-3180
Chen, P. S., Jr., Toribara, T. Y. & Warner, H. (1956)
Anal. Chem. 28, 1756-1758
DePierre, J. W. & Karnovsky, M. L. (1974a) J. Biol.
Chem. 249, 7111-7120
DePierre, J. W. & Karnovsky, M. L. (1974b) J. Biol.
Chem. 249, 7121-7129
Doss, R. C., Carraway, C. A. C. & Carraway, K. L.
(1979) Biochim. Biophys. Acta 570, 96-106
1980
Mammary-tumour ecto-ATPase
Drickamer, K. L. (1975)J. Biol. Chem. 250, 1952-1954
Jarett, L. & Smith, R. M. (1974) J. Biol. Chem. 249,
5195-5199
Jones, B. M. (1966) Nature (London) 212, 362-365
Knight, V. A., Jones, B. M. & Jones, P. C. T. (1966)
Nature (London) 210, 1008-1010
Kruger, P. G., Diamant, B. & Dahlquist, R. (1974) Int.
Arch. Allergy Appl. Immunol. 46, 676-688
Lowry, 0. H., Rosebrough, N. J., Farr, A. L. & Randall,
R. J. (195 1) J. Biol. Chem. 193, 265-275
Luly, P. & Emmelot, P. (1975) Chem.-Biol. Interact. 11,
377-385
Mastro, A. M. & Rozengurt, E. (1976) J. Biol. Chem.
251, 7899-7906
Neilands, J. (1955) Methods Enzymol. 1, 449-454
Novogrodsky, A. (1972) Biochim. Biophys. Acta 266,
343-349
Parkinson, D. K. & Radde, I. C. (1971) Biochim.
Biophys. Acta 242, 238-246
Vol. 191
51
Pommier, G., Ripert, G., Azoulay, E. & Depieds, R.
(1975) Biochim. Biophys. Acta 389,483-494
Riordan, J. R., Slavik, M. & Kartner, N. (1977) J. Biol.
Chem. 252, 5449-5455
Ronquist, G. & Agren, G. K. (1975) Cancer Res. 35,
1402-1406
Rorive, G. & Kleinzeller, A. (1972) Biochim. Biophys.
Acta 274,226-239
Rozengurt, E. & Heppel, L. (1975) Biochem. Biophys.
Res. Commun. 67, 1581-1588
Shin, B. C. & Carraway, K.L. (1973) Biochim. Biophys.
Acta 330, 254-268
Stefanovic, V., Ciesielski-Treska, J., Ebel, A. & Mandel,
F. (1976) Exp. Cell Res. 98, 191-203
Trams, E. G. (1974) Nature (London) 252,480-482
Trams, E. G. & Lauter, C. J. (1974) Biochim. Biophys.
Acta 345, 180-197
Willingham, M. C., Ostland, R. E. & Pastan, I. (1974)
Proc. Natl. Acad. Sci. U.S.A. 71, 4144-4148